Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae

We investigated the succession process of intestinal bacteria during seed production in full-cycle aquaculture of Pacific bluefin tuna (Thunnus orientalis). During the seed production, eggs, healthy fish, rearing water, and feeds from three experimental rounds in 2012 and 2013 were collected before transferring to offshore net cages and subjected to a fragment analysis of the bacterial community structure. We identified a clear succession of intestinal bacteria in bluefin tuna during seed production. While community structures of intestinal bacteria in the early stage of larvae were relatively similar to those of rearing water and feed, the bacterial community structures seen 17 days after hatching were different. Moreover, although intestinal bacteria in the late stage of larvae were less diverse than those in the early stage of larvae, the specific bacteria were predominant, suggesting that the developed intestinal environment of the host puts selection pressure on the bacteria in the late stage. The specific bacteria in the late stage of larvae, which likely composed ‘core microbiota’, were also found on the egg surface. The present study highlights that proper management of the seed production process, including the preparation of rearing water, feeds, and fish eggs, is important for the aquaculture of healthy fish.


Introduction
The intestinal microbiome of mammals has been well studied, and various findings have been obtained. In mammals, infancy and early childhood are the key periods for shaping the gut microbiome [1], which supports host functions such as indirect (immune system-mediated) and direct protection against pathogens [2] and nutrient absorption [3]. Hviid et al. [4] reported that the administration of antibiotics during this key period increases the incidence of other diseases, due to interference with the still unstable gut microbiome. Environmental factors play an important role in shaping the gut microbiome of mammals. The effects of oral acquisition are also important. For example, Ferretti et al. [5] highlighted that oral acquisition of bacteria from maternal skin, breast milk, feces, vagina, and oral cavity is crucial for the Regulations (2021-A-00118). This study was performed in accordance with ARRIVE guidelines. Informed consent was not required for this study. Samples were collected three times: once in 2012 (expressed as Exp12) and twice in 2013 (expressed as Exp13r1 and Exp13r2). We used a full-cycle aquaculture of Pacific bluefin tuna Thunnus orientalis, rearing water, and feeds. The larval fish density was 150,000 fish/30 t tank in Exp12, and 250,000 and 500,000 fishes/50 t tank for Exp13r1 and Exp13r2, respectively. The periods were set from the egg to offshore for the analysis, and the feeding schedule was performed according to Sawada et al. [14] (S1 Fig and S1 Table).
The rearing water and feed were sampled immediately after feeding to the fish, and fish samples were taken immediately before feeding on the next day to investigate the relationship between intestinal and environmental bacteria. The fish samples were collected with a beaker or fry net, depending on the swimming capability. For the feed samples, rotifers (Brachionus plicatilis), Artemia (Artemia salina) nauplii, and feeder larvae (striped beakfish) were placed in a beaker, followed by collection onto a 60-μm-opening nylon membrane, and the commercial pellets were placed in a plastic bag. The rearing water pre-filtered, to remove feeds using a 60μm-opening nylon membrane, was collected into a 50 mL centrifuge tube. All samples were immediately stored at −60˚C until further analysis. The beaker, net, and nylon membranes were washed with hypochlorite.
For the egg and fish samples, we first performed a washing step to remove the bacteria from the samples. Water around the egg (71-120 eggs) and larval (16-44 fish; total length (TL) 2.0-4.6 mm) samples on the nylon membrane were removed with a 47 mm diameter GF/C filter (1822-047, Whatman, Kent, UK; pre-combusted at 450˚C for 2 h) and washed with 5 mL of 0.22-μm-filtered seawater. This washing step was repeated seven times to prevent bacterial contamination from the rearing water. The whole fish body was subjected to DNA extraction because it was difficult to remove only the intestine. For experimental convenience, the bacteria of eggs and small fish were treated as intestinal bacteria. As for the larger larvae and fry (5-44 fish, TL 7.2-65.6 mm), the water around each individual fish was removed in the same manner as described above and washed with 1 mL of 0.22-μm-filtered seawater. This washing step was repeated seven times. The intestines of larvae and fry were removed with a sterilized razor blade and placed into a ZircoPrep Mini tube (FGM50M, NIPPON Genetics Co. Ltd., Tokyo, Japan) containing 400 μL of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0).
For the feed samples, rotifers, Artemia, and the feeder larvae of striped beakfish were washed in the same manner as the egg and fish samples. The weights of rotifer, Artemia, and feeder larvae were 33.2-53.5, 40.5-101.2, and 35.4-82.7 mg, respectively. The commercial pellets were weighed to approximately 50 mg. Each sample was placed in a ZircoPrep Mini tube containing 400 μL of TE buffer. For the water sample, 10 mL of water was filtered through a 0.2-μm-pore size polycarbonate membrane filter (16020002, Advantec Toyo Co. Ltd., Tokyo, Japan) under vacuum < 0.02 MPa. A 0.45-μm-pore size nitrocellulose membrane filter (HAWP02500, Millipore, MA, USA) was used as a base filter to prevent the contamination of filtered seawater. The polycarbonate filter was placed in a ZircoPrep Mini tube containing 400 μL of TE buffer. Filtered seawater was also subjected to filtration in the same manner to check for bacterial contamination.

DNA extraction
The ZircoPrep Mini tubes containing the samples were vortexed for 15 min using a Vortex-Genie 2 (G-560, Scientific Industries, Inc., NY, USA). Lysozyme (final concentration 15 mg/ mL) was added to the tubes and incubated for 60 min at 37˚C with gentle mixing. After incubation, proteinase K (2 mg/mL) and SDS (1.8%) were added and incubated for 60 min at 55˚C with gentle mixing. After incubation, equal amounts of phenol were added to the supernatant and centrifuged at 20,630 × g for 5 min at 20˚C. This phenol step was repeated three times. An equal amount of phenol/chloroform supernatant was added and centrifuged at 20,630 × g for 5 min at 20˚C. Ethanol precipitation was subsequently performed. The DNA was dissolved in 30 μL of TE buffer, and the concentration was measured at 260 nm using a NanoDrop spectrophotometer (ND-1000, Thermo Fisher Scientific Inc., MA, USA).

Automated ribosomal intergenic spacer analysis (ARISA)
PCR amplification was performed in a 20 μL reaction mixture using TaKaRa Ex Taq polymerase (Takara Bio Inc., Shiga, Japan) in a C1000 Thermal Cycler (Bio-Rad, Richmond, CA, USA). The mixture contained 1 × PCR buffer, 2 mM MgCl 2 , 0.2 mM of each dNTP, primers, 0.8 mg/mL bovine serum albumin, and 0.025 units TaKaRa Ex Taq polymerase. The primers used in this study were 16S-1392F (5 -G[C/T]A CAC ACC GCC CA-3´) and 23S-125R (5 -GGG TT[C/G/T] CCC CAT TC[A/G] G-3´) labeled with 6-FAM at the 5´end [24]. The following PCR cycling conditions were used: initial denaturation step at 95˚C for 3 min, 30 cycles of 30 s at 95˚C, 30 s at 56˚C, and 45 s at 72˚C, followed by a final extension step at 72˚C for 7 min. PCR products were analyzed using 2.0% agarose gel electrophoresis.
PCR products were purified using a PCR Clean-Up Mini Kit (Favorgen, Ring-Tung, Taiwan), and the concentration was measured using a NanoDrop spectrophotometer. Purified products were diluted to 25 ng/μL with TE buffer, and 100 ng of a standardized amount were loaded into the fragment analysis. Products were then run for 3 h on an ABI 3130xl Genetic Analyzer (Applied Biosystems, Waltham, MA, USA) with a GeneScan 1200 LIZ dye Size Standard (Applied Biosystems, Waltham, MA, USA). The ARISA peak patterns were analyzed using PeakStudio version 2.2 [25]. According to Chow et al. [26], ARISA peaks were manually binned with maximum bin sizes of 1, 2, 3, and 5 bp for 390-450, 450-650, 650-900, and 900-1200 bp, respectively. Each ARISA peak was regarded as an operational taxonomic unit (OTU), and the relative abundance (peak height divided by the cumulative height of all peaks in the sample) was subjected to community analysis.

Community analysis
All analyses were performed using R software [27]. Diversity indices (OTU number, logarithm of inverse Simpson and Shannon-Weiner indices) were calculated, using the 'Vegan' package [28]. Scatter plots of OTU abundance numbers and diversity indices during experiments were generated and the Pearson's coefficients were calculated using the 'ggpubr' package [29]. Bray-Curtis dissimilarities were calculated, and the distance matrix was analyzed using the between-group average linkage method for clustering and the 'Vegan' package. We performed a similarity profile (SIM-PROF) test (9999 permutations with a p < 0.0001) to determine significant clusters among samples, using the 'clustsig' package [30]. Also, a principal coordinates analysis (PCoA) and an Adonis test (9999 permutations) were used to show OTU variations among samples, using 'ape' [31] and 'Vegan' [28] packages. Venn analysis was performed between the early and late stages of larvae as the result of cluster analysis, using the 'VennDiagram' package [32], to reveal characteristic OTUs. The derived OTUs and abundance were visualized by a heatmap using the 'tidyr' package [33].

Bacterial diversity
During the experiments, bacterial OTU numbers of rearing water, feeds, and intestinal samples ranged from 7 to 81 OTUs, 2 to 80 OTUs, and 1 to 49 OTUs, respectively (Tables 1-3); however, no PCR products were obtained from some intestinal samples (S1 Table). The number of OTUs in the rearing water sample tended to be smaller before the beginning of feeding, and those of feed samples varied, even though the feed type was the same. The

Discussion
In the present study, we demonstrated the succession process of intestinal bacteria in bluefin tuna, in full-cycle aquaculture for the first time. For the analysis, we used a PCR-based fingerprinting analysis, ARISA, which seemed to detect particularly the dominant bacteria in a sample, with some bias [34]. Kashinskaya et al. [35] used several molecular methods to compare differences in the intestinal bacterial communities of Prussian carp and reported that the PCRbased cloning method successfully appeared to detect dominant bacteria in the samples. It is also reported that there is ecological coherence of bacterial diversity pattern between ARISA and deep sequencing techniques [36][37][38]. Therefore, we intended to see a similar trend using the PCR-based method, ARISA, in this study. The dominant bacteria may have a great impact on host development, as can be seen from the probiotic strategy. The simplification in the present study would be key to revealing the succession process of intestinal microbiota. In addition, bacterial contamination from the body surface might be observed in samples of the smaller larvae due to the handling of the whole fish body. Yoshimizu et al. [39] detected bacterial counts in the range of 10 2 −10 3 colony forming units/g of body weight from the body surface of the sac fry. However, the contamination risk could be significantly low in this study as our washing steps were notably more frequent (at least 10 7 times higher) than those of Yoshimizu et al. [39]. Our experiments yield three major findings: 1) the intestinal bacteria in the early stage of larvae were affected by rearing water and feed, 2) the specific intestinal bacteria in the late stage of larvae became dominant regardless of the diversity of rearing water and This study suggests that the management of rearing water and live feeds, such as rotifers, is important for controlling the bacterial communities in seed production because they affect the intestinal bacteria in the early stage of larvae (Figs 1A and S3A). Other studies have also shown that fish intestinal bacteria are mainly affected by feed [12,[40][41][42][43], water [44,45], and both [39,[46][47][48]. Various types of water used for seed production have different bacterial communities [49]. They mentioned that the management of various waters in seed production was a key point because pathogenic microbes would increase during larval development even though the initial rearing water was free of pathogens. Effective management of rearing water has been shown to be 'green-water', a type of rearing water containing a large amount of microalga [50][51][52][53]. Studies have shown that live microalgae such as Nannochloropsis oculata and Chlorella vulgaris could eradicate the pathogenic bacterium Vibrio anguillarum by collaborating with the indigenous probiotic bacterium Sulfitobacter in rearing water. This 'green-water' technique is necessary to maintain the bacterial communities in desirable condition in the early stage of larvae.
Specific bacteria in the late stage of larvae in this study would colonize the favorable intestinal environment of bluefin tuna (Figs 1B and S3B) by the host and/or interactions of hostmicrobe and microbe-microbe [39,[46][47][48][54][55][56]. A pioneering study of the succession process of intestinal bacteria by Yoshimizu et al. [39] indicated the possibility of regulation by the host. They reported that intestinal bacteria in the early stage of salmonid larvae were affected by the rearing water and diet, and then the developed intestinal environment by the activation of digestive tract would establish the characteristic intestinal microbiota. Ontogenetic development of the digestive tract and enzyme activity in bluefin tuna has been reported to develop relatively fast [57,58]. In bluefin tuna farmed at Kindai University, Miyashita et al. [59] showed that activities of pepsin-like and trypsin-like enzymes increased simultaneously with the development of stomach and pyloric caecum functions during the transitional period of juvenile tuna (17)(18)(19)(20)(21)(22)(23)(24)(25) as the rate of percentage of preanal length to standard length increased. The selection pressure would be due to host regulation as well as host-microbe and microbemicrobe interactions [55]. Changes in microbiota contribute to larval health and growth by improving host abilities.
The possible source of characteristic intestinal bacteria in the late stage of larvae would be the egg surface where these characteristic bacteria were also found (Fig 3), and the bacteria should compose of 'core microbiota' [60,61] in bluefin tuna. Fish eggs contain many bacteria on their surfaces [39,41,62], although sometimes the associated bacteria can cause mortality in marine fish [63]. Lauzon et al. [64] demonstrated that probiotic bathing treatment of eggs resulted in the establishment of added bacteria in the larval intestine, leading to increased survival, stress tolerance, and growth of host larvae. Notably, they highlighted that bacterial control was mainly evidenced prior to larval feeding, suggesting the importance of bacterial establishment on the egg surface. The mechanism of transfer from egg surface to larva is still unknown in bluefin tuna, but one possibility is ingestion of egg surface bacteria via grazing on egg debris and/or drinking seawater [62,65]. In the present study, we demonstrated that bacteria on the egg surface would be an important key species for determining 'core microbiota' of intestine in bluefin tuna. As bacteria associated with fish egg surface are likely to be opportunistic species [63], the management of eggs shortly after release, including disinfection [15] and preparation of rearing water, would be an important process. Further investigation is needed to determine whether the key species are still colonized in the adult bluefin tuna.
In conclusion, a clear succession process of intestinal bacteria in full-cycle aquaculture of Pacific bluefin tuna was shown for a total of three times of seed production. While bacteria in the early stage of larvae were affected by bacteria of rearing water and feeds, in the late stage (especially, after 17 DAH when the intestinal environment was developed), they changed to specific bacteria originating from the egg surface, which are likely to be composed of the 'core microbiota'. Thus, we highlighted the importance of proper management in the seed production process, including egg management for the aquaculture of healthy fish. Further research is needed to determine the bacterial function and whether the bacteria are well colonized in adult fish.